Bioconversion of olive mill wastewater into high-added value products

Journal of Cleaner Production 139 (2016) 957e969

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Bioconversion of olive mill wastewater into high-added value products Marianna Dourou a, Anna Kancelista a, 1, Piotr Juszczyk a, 1, Dimitris Sarris b, Stamatia Bellou a, Irene-Eva Triantaphyllidou a, Anita Rywinska a, 1, Seraphim Papanikolaou b, George Aggelis a, * a b

Unit of Microbiology, Division of Genetics, Cell and Development Biology, Department of Biology, University of Patras, Patras, Greece Laboratory of Food Microbiology & Biotechnology, Department of Food Science and Human Nutrition, Agricultural University of Athens, Athens, Greece

a r t i c l e i n f o

a b s t r a c t

Article history: Received 12 May 2016 Received in revised form 27 July 2016 Accepted 26 August 2016 Available online 28 August 2016

Olive mill wastewater (OMW) contains a variety of assimilable carbon sources and therefore it can be regarded as fermentation medium for the production of high-added value products rather than as a waste material. In this study, OMW enriched with low cost carbon sources was used as culture medium for selected yeast strains cultivated in flasks and bioreactors. Lipomyces starkeyi NRRL Y- 11557 and Yarrowia lipolytica strains showed a noteworthy ability to accumulate lipids (15e25%, w/w) cultivated on OMW based media. Oleic acid was the major fatty acid in the lipids produced by the above mentioned yeasts followed by palmitic acid. On OMW media enriched with glycerol Y. lipolytica A6 strain produced also mannitol in considerable amounts (i.e. 13.4 g/L), while Y. lipolytica LGAM S (7) strain produced high quantities of citric acid (i.e. 30.3 g/L). Candida tropicalis LFMB 16 and Saccharomyces cerevisiae MAK-1, cultivated under non-aseptic conditions on OMW media enriched with glucose produced 21.9 and 31.3 g/L of ethanol, respectively. Remarkable phenolic removal was performed by strains of Y. lipolytica, C. tropicalis and S. cerevisiae, while color removal was only observed in trials performed with C. tropicalis and S. cerevisiae. The present study provides a new perspective for the production of high-added value metabolites having biotechnological interest from OMW based media along with OMW management. © 2016 Elsevier Ltd. All rights reserved.

Keywords: Olive mill wastewater Candida tropicalis Lipomyces starkeyi Yarrowia lipolytica Saccharomyces cerevisiae Single cell oil Citric acid Ethanol Mannitol

1. Introduction Olive mill wastewater (OMW) is a dark colored emulsified effluent consisted of the soft pulp tissue of olive fruits, plant and processing waters. Its composition presents a large diversity depending on various parameters such as the variety of olives and their maturity, the region of origin, and especially the technology used for oil extraction (Roig et al., 2006). Three different processes for oil extraction are widely used, namely the traditional pressing processes and two more based on centrifugation, i.e. the three- and two-phase decanting process. The kind and the quantity of the byproducts produced during the olive fruit treatment/oil extraction largely depend on the technology used. Specifically, the traditional

* Corresponding author. E-mail address: [email protected] (G. Aggelis). 1 Current address: Department of Biotechnology and Food Microbiology, Wroclaw University of Environmental and Life Sciences, Wroclaw, Poland. http://dx.doi.org/10.1016/j.jclepro.2016.08.133 0959-6526/© 2016 Elsevier Ltd. All rights reserved.

methods and the three-phase decanting process produce, besides olive oil, a liquid extract (namely OMW) and a solid residue (namely olive pomace) as final byproducts. On the other hand, in the twophase decanting process significantly less water is required and the only byproduct is a sludge-like pomace (namely olive wet pomace). This type of process has been proposed as an “eco” alternative way, environmentally friendlier comparing to the threephase decanting processes and for this reason is widely used in the last few years, although the viscous residue, which is produced in significant amounts, is a hard to manage polluting waste (Salomone and Ioppolo, 2012; Therios, 2009). The major environmental problems associated with olive oil extraction mills are related in both the large volumes of water required and the ineffective management of OMW and olive pomace. Commonly OMW is characterized by high chemical and biological oxygen demand and contains organic components such as carbohydrates, polysaccharides, fatty acids, polyalcohols, pectins, tannins and phenolic compounds (Lesage-Meessen et al., 2001), rendering this residue a severe pollutant mainly for Mediterranean

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countries in which olive oil production accounts for more than 90% of the world production (McNamara et al., 2008). Additionally, the large volume of OMW produced (i.e. up to 30
106 m3 per year), in combination with the short discarding time, which is usually defined from October to February, increases the importance of this waste from both an environmental and financial point of view (Crognale et al., 2006). The olive oil industry is one of the most important food industries in many countries in the Mediterranean area. Specifically, in Greece, olive cultivation dates back more than 4.000 years ago, while the olive fruit and oil manufacturing sector plays a relevant role in the economic development of this country. Greece is the world's third largest producer of olive oil, coming after Spain and Italy and before Tunisia and Portugal. Olive trees in Greece grow mainly on hilly land and geographically speaking, almost 80% of olive oil production is centered in three regions: Peloponnese (37%), Crete (30%) and the Ionian Islands (12%). Between 2000/01 and 2009/10, Greek production of olive oil ranged from a low of 305.000 t (2008/09) to a record high of 435.000 t (2004/05), averaging out at 366 150 t/year, according to the International Olive Oil Council (IOOC, 2012). When disposed into the environment, OMW creates serious environmental problems. For instance, its discarding into watercourses could lead to deterioration of natural water bodies that is a serious threat to the aquatic life (Karaouzas et al., 2011). Pollution of ground and surface waters, soil contamination, odor nuisance, environmental degradation, as well as, effects of toxicity and growth inhibition on species from different trophic levels have been reported (Aggelis et al., 2003). Currently, the methods applied for OMW treatment are either physico-chemical or biological. Physicochemical methods, including simple evaporation, reverse osmosis, ultrafiltration, coagulation, oxidation, thermal drying and advanced oxidation processes (i.e. ozonation, Fenton process, electrochemical oxidizing methods) (Cayuela et al., 2006; Ena et al., 2012; Khoufi et al., 2006; Mert et al., 2010; Sarika et al., 2005; Scoma et al., 2011), are of high cost, so few of them have been applied on an industrial scale. For instance, olive pomace is treated via thermal drying as fuel for electric and thermal energy production, but the economic feasibility seems to be low. On the other hand, anaerobic digestion has been proposed as a promising technology for olive pomace management and energy (i.e. biogas) production, although researchers have to overcome many problems, such as growth inhibition of methanogens from phenolic compounds, low pH, alkalinity, low nitrogen content and many others (Orive et al., 2016). Other biological methods, such as aerobic treatment, composting, vermicomposting together with other agro-industrial residues (Chowdhury et al., 2015, 2013; Dajko and Vasilikiotis, 2015; Dhouib et al., 2006; Tortosa et al., 2012; Tziotzios et al., 2007) have also been suggested, but the obtained results are rather mediocre. OMW spreading on the arable land of agricultural regions is a promising recycling method, since OMW, containing nutrients and inducing beneficial modifications in soil structure, may have, under certain circumstances, favorable effects on plant growth (Altieri and Esposito, 2010; El Hassani et al., 2010). Furthermore, as the research on natural dyes is constantly increasing, Meksi et al. (2012) proposed OMW as a potential source for textile dyeing, as it contains a valuable potential of abundant natural coloring substances. The research on OMW biotreatment focuses on the degradation of phenolic compounds, since these compounds are principally responsible for phytotoxicity and microbial growth inhibition (Assas et al., 2002; Casa et al., 2003; Ergül et al., 2009; Sampedro et al., 2009; Sarris et al., 2014, 2013), while their breakdown is considered to be the limiting step during biotreatment. In addition, phenolic compounds are a serious threat to human health, due to

their carcinogenic properties and therefore, phenol has been registered as priority pollutant by the US Environmental Protection Agency with a tolerable limit of 0.1 mg/L in wastewaters (USEPA, 1991). Several biological processes using microorganisms belonging to the genera Candida, Yarrowia, Pleurotus, Geotrichum, Lentinus, Phanerochaete, Pseudomonas, Sphingomonas, Ralstonia, Rhodotorula etc. have been proposed (Aggelis et al., 2003; Amaral et al., 2012; Assas et al., 2002; D'Annibale et al., 1998; Dhouib et al., 2006; Di Gioia et al., 2001; Dias et al., 2004; Fadil et al., 2003; García et al., 2000; Karakaya et al., 2012; Kissi et al., 2001; Ntougias et al., 2012; Sayadi et al., 2000; Scioli and Vollaro, 1997; Tsioulpas et al., 2002). However, most of them have been proved expensive and/or rather ineffective, leading to the accumulation of toxic condensed tannins and polyphenolics of high molecular weight (Aggelis et al., 2003; Assas et al., 2002; Hamdi, 1993; Sayadi et al., 2000). From a biotechnological standpoint OMW, containing various carbon sources, organic compounds and minerals, would be considered as an exploitable resource that could be valorized as substrate in various bioprocesses (Lanciotti et al., 2005). Actually, several microorganisms have been proved able to grow on OMW and produce various high-added value products (i.e. enzymes, organic acids, exopolysaccharides, single cell oils-SCOs, etc.) (Arous et al., 2016; Bellou et al., 2014; D'Annibale et al., 2006; Mateo and Maicas, 2014; Papanikolaou et al., 2008; Sarris et al., 2014, 2013, 2011; Scioli and Vollaro, 1997; Yousuf et al., 2010). Wood-rotting fungus, such as Pleurotus species, Lentinula edodes and Hericium erinaceus, have drawn research attention due to their culinary value and their content in bioactive compounds with beneficial effects in health (i.e. antioxidants, vitamins, b-glucans, and lectins). Few investigations report bioconversion of OMW into high-quality biomass (Koutrotsios et al., 2016; Lakhtar et al., 2010; Zervakis et al., 2013), in parallel with OMW detoxification. The aim of the present research was to further investigate the ability of technologically suitable yeasts, such as Candida tropicalis, Lipomyces starkeyi, Saccharomyces cerevisiae and Yarrowia lipolytica, to grow on OMW (potentially enriched with other carbon sources of low acquisition cost) and produce high-added value metabolites, such as SCOs, citric acid, ethanol and mannitol, with concurrent degradation of phenolic compounds. The strategy followed in order to reach this aim is depicted in Fig. 1. It was found that on OMW based media L. starkeyi and Y. lipolytica A6 and S11 strains were able to accumulate lipids, while A6 produced also mannitol. Y. lipolytica LGAM S (7) strain produced high quantities of citric acid, while C. tropicalis and S. cerevisiae were able to produce ethanol, simultaneously provoking remarkable phenolic and color removal. 2. Materials and methods 2.1. Microorganisms and culture conditions The following yeast strains were used: Candida tropicalis LFMB 16 (culture collection of the Agricultural University of Athens, Greece), Cryptococcus curvatus NRRL Y- 1511, Lipomyces lipofer NRRL Y- 11555, L. starkeyi NRRL Y- 11557, L. tetrasporus NRRL Y- 11562, Saccharomyces cerevisiae MAK-1 (National Agricultural Research Foundation, Athens, Greece) and Yarrowia lipolytica strain LGAM S (7) (culture collection of Agricultural University of Athens, Greece) and strains A6, S5, S6, S9-11 and S17 (Wroclaw University of Environmental and Life Science, Poland). The strains were maintained on potato dextrose agar (PDA, Himedia, India) at 4 ± 1  C and regularly sub-cultured. The culture media consisted of OMW (obtained from an olive oil three-phase decanter manufacture located in Patras industrial zone, Western Greece) enriched with carbon sources and minerals

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Fig. 1. Process flow chart for OMW conversion into microbial products. The final products were characterized according to the colors of biotechnology (BT): Yellow, Food BT; Green, Environmental BT; Red, Pharmaceutical BT; Pink, Chemical Industry. Grey was used for characterization of waste products. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

(see below). The OMW solid particles were removed after filtration and subsequent centrifugation at 15.000 rpm, 4  C for 30 min. The supernatant, diluted in various rates with tap water, was enriched with commercial glucose (95% w/w purity, Hellenic Industry of Sugar S.A., Thessaloniki, Greece) or glycerol (98% purity, Fluka, Steinheim, Germany), added in various concentrations, and with the following minerals (in g/L): KH2PO4 (Fluka), 12.0; Na2PO4 (Fluka), 12.0; MgSO4.7H2O (Fluka), 1.5; CaCl2.2H2O (Carlo Erba, Rodano, Italy), 0.1; ZnSO4.7H2O (Merck), 0.001; MnSO4.5H2O (Fluka), 0.0001; CuSO4.5H2O (BDH), 0.0001; Co(NO3).3H2O (Merck), 0.0001. (NH4)2SO4 (Carlo Erba) at 0.5 g/L and yeast extract (Conda, Madrid, Spain) at various concentrations (i.e. 0.5, 1.0 or 3.0 g/L), were used as nitrogen sources. Flask cultures were conducted in 250 mL Erlenmeyer flasks containing 50 ± 1 mL of growth medium. After sterilization (at 121  C for 20 min) the flasks were inoculated with 1 mL (108 cells) of a pre-culture carried out on potato dextrose broth, and incubated

in a rotary shaker (ZHICHENG ZHWY 211C, Shanghai, China) at temperature 28 ± 1  C and agitation rate 180 rpm. The medium pH was maintained at desirable levels (i.e. 2.8, 3.5, 4.5, 6 ± 0.03) by adding, if needed, 4 M NaOH (Merck) or 4 M HCL (Sigma, Steinheim, Germany) solutions under aseptic conditions. Bioreactor cultures were performed under aseptic conditions in a laboratory bioreactor (Bioengineering, Ralf Plus-System, Wald ZH, Switzerland) of total volume 3.7 L and working volume 1.6 L. The reactor was sterilized at 121  C for 120 min and kept at room temperature for 48 h to ensure medium sterility. The culture vessel was inoculated with 200 mL of a pre-culture prepared as described above. Dissolved oxygen concentration was kept at 20% of saturation value through a cascade controlling incoming gas (mixtures of air and pure oxygen) flow rate and composition. In all cases, incoming gas passed through a bacteriological filter of 0.2 mm pore size (Whatman). Medium pH was automatically controlled at 6 ± 0.03 by adding 1 M NaOH (Merck) solution. Agitation rate was

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280 rpm and the incubation temperature was controlled at 28 ± 0.1  C. Antifoam A (Fluka) was added if necessary. Non-aseptic cultures were performed in a laboratory scale bioreactor (MBR, AG Switzerland) of total volume 3.5 L and working volume 3.0 L as described by Sarris et al. (2014, 2013). Cultures under non-aseptic conditions were also carried out in a laboratory home-made glass bioreactor, resembling to an open pond, of total volume 8.7 L and working volume 5 L (Bellou and Aggelis, 2012). Glass bioreactor was operating at room temperature (25 ± 3  C) and pH was controlled at 2.8 ± 0.03 by adding 4 M HCL solution. The culture was inoculated with 500 mL of a pre-culture prepared as above.

2.2. Biomass determination and analytical methods Biomass was harvested by centrifugation at 15.000 rpm for 15 min at 4  C using a Heraeus (Biofuge Stratus, Osterode, Germany) centrifuge. Cells were washed three times with NaCl 0.9% and small amount of hexane (for extracellular lipids removal) during centrifugations. Dry biomass (X, g/L) was determined gravimetrically after drying at 80e100  C until constant weight. The supernatant was collected and stored at 20  C for further analysis. Glycerol, citric acid and ethanol were determined in filtered aliquots of the culture supernatant by HPLC (Ultimate 3000 Dionex, Germering, Germany) equipped with RI and UV-Vis detectors and an Aminex HPX-87H column (300 mm
7.8 mm) as described in Makri et al. (2010) and Sarris et al. (2013). Mannitol was determined according to Tomaszewska et al. (2012), using Carbohydrate H þ Column (Thermo Scientific, Waltham, MA). Reducing sugar concentration in the growth medium was determined according to Miller (1959) and expressed as glucose (g/ L). The total phenolic compounds were determined according to Folin and Ciocalteau (1927) and expressed as gallic acid equivalent. Decolorization of the medium was measured at 395 nm. Substrate lipids (i.e. those contained in OMW) were extracted using hexane as solvent after medium acidification with equal volume of a 4 M HCl solution. The organic phase was collected, dried over anhydrous Na2SO4 (Sigma) and the lipids were gravimetrically determined after solvent evaporation under vacuum. Yeast lipid extraction was performed according to Folch et al. (1957) with slight modifications as described by Bellou et al. (2014). Fractionation of total lipids into neutral (NL), glycolipids plus sphingolipids (G þ S) and phospholipids (P) was performed by silicic acid chromatography. Lipids (approximately 100 mg) were dissolved in 1 mL chloroform (Fluka), and fractionated by using a column (25 mm
100 mm) containing 1 g silicic acid (Fluka), activated by heating overnight at 80  C. Successive applications of dichloromethane (Sigma) 100 mL, acetone (Fluka) 100 mL and methanol (Sigma) 50 mL, produced fractions containing NL, G þ S and P, respectively (Fakas et al., 2007). After evaporation of the respective solvent, lipid fractions were collected and stored under argon atmosphere at 20  C. Fatty acid composition of cellular or substrate lipids and of each lipid fraction was performed by gas chromatography (GC) after trans-methylation, according to the AFNOR (1984) method. GC apparatus (Agilent Technologies 7890 A) was equipped with a flame ionization detector and a HP-88 (60 m
0.25 mm) column (J&W scientific). GC conditions were as follows. Carrier gas: helium; flow rate: 1 mL/min; injection temperature 250  C; oven temperature 200  C; FID temperature 280  C. Peaks of methyl esters were identified by reference to authentic standards. Visualization of intracellular lipid droplets was performed by fluorescence microscopy after staining yeast cells with Nile Red (Greenspan, 1985).

3. Results and discussion 3.1. Single cell oil and mannitol production Oleaginous microorganisms have the ability to assimilate a wide range of low-value argo-industrial residues, such as raw glycerol, OMW and several other by-products. Using such residues as substrates the production cost of SCO can be reduced and beneficial effects for the environment can be arisen. In this study, the ability of Cryptococcus curvatus, Lipomyces lipofer, L. starkeyi and L. tetrasporus to produce SCO was tested on media containing OMW diluted in water, so as to obtain an initial phenolic compound concentration around 2 g/L, and enriched with a) glycerol 50 g/L or b) glucose 30 g/L. Media containing phenolic compounds in higher concentrations inhibited yeast growth. Although all strains were able to grow in the presence of OMW, L. starkeyi showed an improved capacity to grow and accumulate lipids and therefore it was selected for further studies. Also, preliminary studies with Y. lipolytica strains able to grow at low pH were conducted on OMW media containing glucose 40 g/L and two strains (i.e. A6 and S11) were selected for further studies. L. starkeyi is an interesting yeast regarded as a non-conventional source of oil (Xavier and Franco, 2014; Yousuf et al., 2010). When OMW was used as sole carbon and energy source, the growth of L. starkeyi was mediocre, since only 2.7 g/L of dry biomass (Xmax) were synthesized (Fig. 2a). However non-negligible lipid quantities (i.e. L/Xmax ¼ 17.8% w/w) were accumulated within the yeast cells. When OMW was enriched with 30 g/L of glucose, L. starkeyi produced 9.5 g/L of Xmax containing 25.4% w/w lipids (Fig. 2b). It seems that the presence of OMW in the growth medium negatively affected lipid accumulation but not growth of L. starkeyi, since when the strain was cultivated on glucose 30 g/L in absence of OMW, 10.9 g/L of Xmax were produced, but the lipid content was much higher (L/Xmax ¼ 34.5%) (Fig. S1). Similarly, the conversion yield of the carbon substrate to lipid (YL/S) was 0.07 g/g in the presence of OMW and 0.11 g/g in the absence of OMW, indicating again the negative effect of OMW on the liposynthetic machinery of L. starkeyi. This is in disagreement with Bellou et al. (2014) and Sarris et al. (2011) in which OMW has been considered as a “lipogenic” medium. On the other hand, concerning the maximum value of biomass yield per unit of substrate (YX/S), the highest value obtained for the OMW fermentation (i.e. 0.39 g/g) was similar to those obtained from OMW plus glucose and glucose fermentation (0.37 and 0.33 g/g, respectively), suggesting that the growth of L. starkeyi was essentially unaffected by the presence of the OMW. In Fig. 3, the accumulated lipids within the yeast cells are depicted in the form of lipid droplets, after staining with Nile Red. In all the above mentioned cases satisfactory substrate (reducing sugars) consumption was observed. Several authors reported that L. starkeyi showed efficiency in assimilation of a variety of carbon sources such as glucose, xylose and hemicellulosic hydrolysate (El-Naggar et al., 2011; Sha, 2013; Wild et al., 2010; Yousuf et al., 2010), being able in valorizing a variety of agro-industrial by-products. However, it appears that under the above mentioned culture conditions L. starkeyi was not able to remove phenolic compounds of OMW, while no decolorization of the medium was observed (data not shown). Considering that OMW is a very complex effluent, Yousuf et al. (2010) investigated some parameters (composition, pH, temperature, etc.) of OMW that affect the growth of L. starkeyi. Apart from the phenolic compounds, OMW fatty acids (even found in low concentrations in the medium, i.e. 0.52 g/L) should be taken into account, since their presence may limit the efficiency of the processes. According to the current investigation the various substrate fatty acids were selectively taken up by the yeast cells. Specifically, a selective uptake of oleic acid (D9C18:1) (the dominant

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Fig. 2. Kinetics of growth, lipid accumulation and substrate consumption of Lipomyces starkeyi cultivated in flasks on media containing OMW (a); OMW enriched with glucose 30 g/ L (b). Culture conditions: pH 6 ± 0.03; Initial phenolic compounds 1.9 g/L; T ¼ 28  C. Abbreviations: X, dry biomass; L/X%, lipids in dry biomass %; R.S., reducing sugars.

fatty acid in OMW lipids) was observed, regardless the enrichment of the growth medium with other carbon sources, as its concentration in the extracellular lipids decreased with the time (data not shown). D9C18:1 was the most abundant fatty acid in L. starkeyi lipids, representing 50e63% of total fatty acids (Table 1). D9C18:1 concentration showed no remarkable change during OMW fermentation. In contrast, on OMW plus glucose or glucose fermentation its concentration was constantly decreased with time. Palmitic (C16:0) and linoleic (D9,12C18:2) acids were also found in considerable concentrations in L. starkeyi lipids, regardless of the growth medium. Specifically, on OMW media the concentration of C16:0 in the cellular lipids decreased with time, while, on the contrary, on OMW plus glucose or solely glucose media, increased. These results are in agreement with those reported in El-Naggar et al. (2011), Sha (2013), Wild et al. (2010), Xavier and Franco (2014) and Yousuf et al. (2010). Additionally, Wild et al. (2010) reported that the nature of the carbon substrate affected the fatty acid

composition of the yeast lipids. Likewise, Angerbauer et al. (2008) reported that small amounts of myristic (C14:0), linolenic (C18:3), arachidic (C20:0), gadoleic (C20:1) and behenic (C22:0) acids were also present in the lipids produced by L. starkeyi grown on sewage sludge. Low pH media are suitable for large scale applications since they protect the culture against bacterial contamination. Indeed, in some very low pH media sterilization can be omitted and thus the fermentation cost can be significantly reduced. In the present investigation the acid tolerant strain A6 of Yarrowia lipolytica was cultivated at pH 2.8. When cultures were carried out on OMW without carbon source supplementation, Xmax did not exceed 2.2 g/ L and L/Xmax% reached the value of 19.1% w/w, while when the same medium was enriched with glycerol (50 g/L), 5.6 g/L of Xmax containing 14.9% lipids were produced (Table 2a). In the latter case glycerol was preferentially taken up, while OMW reducing sugars were discriminated. Remarkably, under these culture conditions,

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Fig. 3. Morphology of yeast cells and lipid droplets of Lipomyces starkeyi cultivated on OMW (a); OMW enriched with glucose 30 g/L (b); glucose 30 g/L (c), under optical (top pictures) and fluorescence (bottom pictures) microscope after staining with Nile Red. Magnification: 1000
.

Table 1 Fatty acid composition (%, w/w) of lipids produced during different growth phases of Lipomyces starkeyi, Yarrowia lipolytica (strains A6, S11, LGAM S 7) on OMW, OMW plus glucose, OMW plus glycerol or glucose. Data are presented as mean values from duplicate experiments. Culture conditions: as described in the text. Abbreviations: F.C., flask culture; B.C., bioreactor culture. Exponential growth phase: At fermentation time 18e50 h; Early stationary growth phase: 50e100 h; Late stationary phase: 100e220 h Strain

Medium

Type of culture

Growth phase

C16:0

L. starkeyi NRRL Y- 11557

OMW

F.C.

OMW þ Glucose 30 g/L

F.C.

Glucose 30 g/L

F.C.

OMW þ Glycerol 50 g/L

F.C.

OMW þ Glycerol 50 g/L

B.C.

Glucose 40 g/L

F.C.

OMW þ Glucose 50 g/L

F.C.

Glucose 40 g/L

F.C.

OMW

F.C.

OMW þ Glycerol 50 g/L

F.C.

Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary Exponential Early stationary Late stationary

21.2 22.5 15.0 19.2 33.7 31.0 31.5 37.7 37.2 12.1 12.5 12.5 14.9 15.7 13.2 15.2 14.2 12.1 12.7 12.4 12.8 14.8 14.3 11.7 16.4 12.5 10.9 12.7 13.8 16.3

Y. lipolytica A6

Y. lipolytica S11

Y. lipolytica LGAM S (7)

the strain produced mannitol in significant amounts, i.e. 13.4 g/L after 216 h of cultivation (Fig. S2). Mannitol is a sugar widely used in the food, pharmaceutical and chemical industries. Although it can be produced by chemical synthesis, its microbial production is much more attractive (Saha and Racine, 2011; Tomaszewska et al., 2012), while Y. lipolytica strains are potential candidates for large scale applications (Tomaszewska et al., 2012). The ability of

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

D9

C18:0

D9

2.1 ± 0.3 3.3 ± 0.5 2.5 ± 0.2 2.1 ± 0.3 3.5 ± 0.2 3.6 ± 0.4 3.3 ± 0.5 3.4 ± 0.6 5.0 ± 0.5 5.6 ± 0.6 7.0 ± 0.6 8.3 ± 0.1 9.3 ± 0.6 8.6 ± 0.9 8.4 ± 0.6 8.2 ± 0.7 11.7 ± 0.8 14.0 ± 0.4 11.0 ± 0.3 11.4 ± 0.7 12.6 ± 1.5 9.4 ± 0.2 16.5 ± 1.1 18.3 ± 1.0 7.2 ± 1.3 9.0 ± 0.1 6.2 ± 0.1 4.8 ± 1.2 7.7 ± 0.9 11.1 ± 1.1

3.7 ± 0.5 2.7 ± 0.1 2.2 ± 0.06 3.6 ± 0.2 5.2 ± 0.8 4.4 ± 0.4 4.0 ± 1.9 7.1 ± 0.8 6.8 ± 0.5 6.7 ± 1.1 8.4 ± 0.3 8.1 ± 0.3 16.6 ± 3.8 16.7 ± 0.6 14.4 ± 0.7 10.3 ± 2.0 7.2 ± 0.4 6.1 ± 0.2 5.8 ± 1.0 8.2 ± 0.2 7.9 ± 0.5 7.5 ± 0.0 7.5 ± 1.2 6.5 ± 0.3 5.2 ± 1.2 4.9 ± 0.5 5.2 ± 0.5 6.2 ± 0.6 8.7 ± 2.0 8.5 ± 0.6

63.5 58.1 67.8 65.2 52.2 51.2 50.4 47.2 46.2 60.1 54.5 52.0 56.5 50.6 54.6 58.9 57.0 55.7 59.0 57.6 55.8 59.7 56.0 55.2 52.1 47.1 42.4 58.6 55.8 45.1

C16:1

0.3 2.5 1.5 1.9 1.9 0.7 2.0 1.2 0.8 0.2 0.3 0.3 1.2 0.6 0.3 0.4 0.9 0.3 0.5 0.6 0.7 0.8 0.1 0.4 0.6 1.8 0.2 1.2 1.2 1.0

C18:1 ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

2.4 1.7 1.0 2.9 0.3 0.6 4.8 1.1 0.2 2.5 0.9 0.0 2.7 3.4 2.8 0.5 0.4 0.08 0.8 2.0 2.5 0.2 0.8 1.2 1.5 1.2 0.6 1.5 2.2 2.1

D9,12

C18:2

11.6 ± 1.1 11.1 ± 0.5 9.5 ± 0.4 8.2 ± 0.2 3.2 ± 0.7 2.5 ± 0.5 8.1 ± 4.6 2.4 ± 0.3 4.5 ± 0.6 13.6 ± 0.9 15.4 ± 0.5 17.3 ± 0.6 5.0 ± 0.3 3.9 ± 0.6 4.6 ± 0.1 7.8 ± 0.4 9.7 ± 0.9 11.3 ± 0.5 9.9 ± 1.3 9.7 ± 1.7 11.5 ± 0.5 7.4 ± 0.9 6.9 ± 1.0 7.0 ± 0.3 20.1 ± 1.2 21.1 ± 0.2 19.4 ± 0.5 16.9 ± 1.0 16.0 ± 0.5 16.5 ± 1.1

Y. lipolytica to produce mannitol on OMW based media is of significance for the valorization of the effluent. When the strain was cultivated under non-aseptic conditions in an open pond-like home-made glass bioreactor on OMW enriched with glycerol (50 g/L) (Fig. S3), Xmax was unaffected, while L/Xmax% reached the value of 11.4% w/w in dry biomass (Fig. 4). Under these culture conditions, lipid degradation was also observed. It seems that some

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Table 2 Growth and consumed reducing sugars at the fermentation time in which maximum lipid accumulation was achieved by Yarrowia lipolytica A6 (a) and Yarrowia lipolytica S11 (b) cultivated in flasks on OMW, OMW enriched with glycerol or glucose and on glucose. Culture conditions: pH 2.8 ± 0.03; Initial phenolic compounds 1.9 g/L; T ¼ 28  C. Abbreviations: as in Fig. 1. Strain

Medium

t (h)

L/X%max

(a)

Y. lipolytica A6

(b)

Y. lipolytica S11

OMW OMW þ Glycerol 50 g/L Glucose 40 g/L OMW þ Glucose 50 g/L Glucose 40 g/L

118 118 118 118 120

19.1 14.9 20.5 16.5 24.7

± ± ± ± ±

0.04 1.9 0.3 0.1 0.4

X (g/L) 1.5 5.6 7.2 5.4 7.3

± ± ± ± ±

0.1 0.4 0.1 0.2 0.4

Consumed R.S. (g/L) 5.9 ± 0.03 1.1 ± 0.3 35.4 ± 0.9 37.4 ± 0.5 39.3 ± 0.2

Fig. 4. Kinetics of growth, lipid accumulation and substrate consumption of Yarrowia lipolytica A6 cultivated in bioreactor on media containing OMW enriched with glycerol 50 g/L. Culture conditions: Non-aseptic; pH 2.8 ± 0.03; Initial phenolic compounds 1.9 g/L; T ¼ 25 ± 3  C (room temperature).; Abbreviations: as in Fig. 2; Glol, glycerol.

OMW components negatively affected both growth and lipid accumulation of A6 strain, since when the strain was cultivated on a synthetic medium in flasks (double limited in nitrogen and magnesium) containing glucose 40 g/L as carbon source, Xmax was 7.2 g/ L and L/Xmax% around 21% (Table 2a). D9C18:1 was the dominant fatty acid in the Y. lipolytica A6 lipids, followed by C16:0, C18:0 and D9,12C18:2 acids (Table 1). The fatty acid profile of A6 strain observed in this work was similar to that reported in previous investigations (Makri et al., 2010; Papanikolaou and Aggelis, 2002), although the lipids produced on OMW plus glycerol media were enriched in D9C16:1, and C18:0, but not in D9C18:1, with the time. It seems that the fatty acid composition was affected by the presence of OMW in the growth environment, since in the case of glucose fermentation, the concentration of both C16:0 and C18:0 constantly decreased with time. Y. lipolytica lipid contained higher amounts of NL (74.8%, w/w on total lipid), following by P (18.7%, w/w on total lipid) and G þ S (6.6%, w/w on total lipid). The mono-unsaturated C18:1 was preferentially esterified in NL, while its polyunsaturated homologue C18:2 was the major fatty acid of P. C16:1 was mainly found in G fraction. Another interesting strain able to grow at pH 2.8 is Y. lipolytica S11. According to Juszczyk et al. (2013) this strain was able to efficiently convert raw glycerol into biomass rich in protein of high nutritional value. In this current study, in OMW media enriched with 50 g/L glucose, the above strain produced 5.2 g/L of Xmax (YX/ S ¼ 0.25 g/g) having L/Xmax ¼ 16.5% w/w. It seems again that the presence of OMW negatively affected growth and lipid biosynthesis, since both biomass and lipid production were higher (i.e.

7.4 g/L and 24.7%, respectively), on synthetic media having 40 g/L glucose as carbon source (Table 2b). However, comparing the current results with those reported in Juszczyk et al. (2013) we conclude that S11 strain achieved satisfactory yields on OMW based media. Lipids produced on OMW plus glucose had a similar fatty acid composition to those produced on glucose (Table 1). In both cases, D9C18:1 was found at high percentages, up to 59% in total lipids. C16:0 and D9,12C18:2 were also found in considerable percentages, while their concentration remained invariable with time. However, the concentration of D9C16:1 in total lipids showed remarkable increase with time on glucose, but not on OMW plus glucose media. Lipid content and fatty acid composition is a major criterion characterizing the biomass nutritional value (Juszczyk et al., 2013). Although Y. lipolytica can accumulate high amounts of lipids (more than 30% w/w in dry biomass) (Papanikolaou and Aggelis, 2011, 2002), under the culture conditions mentioned above, lower amounts of lipids were accumulated within the cells. During growth of Y. lipolytica on glucose or similarly metabolized compounds (e.g. glycerol), under conditions that favor the production of lipid (and potentially of citric acid), the yeast may possibly show typical “oleaginous” features, in which normally after nitrogen exhaustion, lipid in significant quantities is accumulated, while negligible quantities of citric acid are secreted in the medium. However, this is probably the scarcest case for Y. lipolytica (Fontanille et al., 2012) since usually after nitrogen exhaustion, lipid initially is stored but shortly, before the exhaustion of the carbon source, is degraded, while citric acid is excreted in the medium

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(Makri et al., 2010; Papanikolaou et al., 2013). Alternatively, lipid is accumulated with slow rates within the yeast cells (but it is not subjected to degradation), while citric acid is constantly being
et al., 2009; Dobrowolski et al., secreted into the medium (Andre 2016). Contrary to L. starkeyi, both A6 and S11 strains of Y. lipolytica exhibited a considerable capacity in phenolic removal, which is considered important for the biological treatment of OMW (Hamdi et al., 1992). The maximum removal (up to 30%) was achieved by Y. lipolytica S11, while Y. lipolytica A6 in bioreactor cultures reduced phenolic compounds by 12e16% (data not shown). Various microorganisms were proved efficient in reducing significantly the concentration of phenolic compounds, such as Geotrichum candidum (Aissam et al., 2007; Assas et al., 2002; García et al., 2000), Pleurotus ostreatus (Aggelis et al., 2003), Aspergillus niger (Crognale et al., 2006; García et al., 2000), Candida boidinii (Aissam et al., 2007) and Saccharomyces cerevisiae (Sarris et al., 2013). Despite remarkable removal of phenolic compounds by A6 and S11 strains, no color removal was observed, although Papanikolaou et al. (2008) and Sarris et al. (2011) reported a remarkable decolorization of OMW using various Y. lipolytica strains. In general, the potential of several yeast strains to decrease phenolics concentration and color in OMW remains confusing. While some strains belonging to Saccharomyces and Yarrowia are capable to significantly reduce color, and to lesser extent remove phenolics (D'Annibale et al., 2006; Lanciotti et al., 2005; Papanikolaou et al., 2008; Sarris et al., 2014, 2013, 2011), some others (De Felice et al., 1997; Scioli and Vollaro, 1997; and also results of the current investigation) seem incompetent to reduce phenolics concentration and color. However, it should be indicated that color intensity is dependent on several physicochemical parameters, thus not proportional to phenolics concentration (Aggelis et al., 2003; Bellou et al., 2014; Sarris et al., 2014, 2013, 2011; Tsioulpas et al., 2002). 3.2. Citric acid production Various Y. lipolytica strains have been reported for their capability to convert several agro-industrial residues into metabolic products such as SCO and organic acids (mainly citric acid) (Finogenova et al., 2005; Makri et al., 2010; Papanikolaou et al.,
ska and Rymowicz, 2011; Rywin
ska et al., 2013). 2002; Rywin However, production of citric acid by microorganisms cultivated on OMW based media has been reported in a scarce number of investigations (Papanikolaou et al., 2008; Sarris et al., 2011), while to the best of our knowledge, the current investigation is the first report in the international literature in which blends of OMW with glycerol have been employed as simultaneous carbon sources and fermentation medium amenable for the production of citric acid. Y. lipolytica LGAM S (7) cultivated on OMW diluted in water (initial phenolic compound concentration around 2 g/L), produced Xmax ¼ 5.2 g/L with L/Xmax ¼ 6.3% (Fig. 5). Only 6.3 g/L of citric acid were produced, which is very low considering that OMW naturally contains 4.7 g/L citric acid. However, when OMW was enriched with 50 g/L glycerol, 30.3 g/L of citric acid were produced. The concomitant conversion yield of citric acid produced per unit of substrate (glycerol þ reducing sugars) consumed (YCA/S) was 0.62 g/ g. Xmax was 9.3 g/L and low amounts of lipids were accumulated within the cells (i.e. L/Xmax ¼ 9.8%) (Fig. 6a, b). Glycerol uptake rate was much higher than the one of the reducing sugars (Fig. 6b). Likewise, Papanikolaou et al. (2016) and Workman et al. (2013) reported for several Y. lipolytica strains grown on media containing both glycerol and glucose as carbon sources that glycerol was more rapidly and efficiently assimilated than glucose, suggesting that the preferential assimilation of glycerol vs glucose is a common feature for Y. lipolytica strains. In bioreactor trials, both citric acid

and Xmax production was lower (21 g/L and 7.9 g/L, respectively), but YCA/S and L/Xmax values higher (0.68 g/g and 15%, respectively) than those observed in flasks (Fig. S4). As for citric acid yield, the results obtained in the current research are similar to those reported in Papanikolaou et al. (2002) for Y. lipolytica cultivated on raw glycerol. However, higher YCA/S (up to 0.90 g/g), and citric acid production (ranging between 50 and 155 g/L) were reported in Kamzolova et al. (2011), Morgunov et al. (2013), Papanikolaou et al.
ska and Rymowicz (2011, (2013), Rymowicz et al. (2006), Rywin
ska et al. (2010). 2010) and Rywin Concerning the substrate uptake, Y. lipolytica LGAM S (7), contrary to the strain A6, was able to consume reducing sugars from OMW, regardless of the presence of other carbon sources (i.e. glycerol). Lipid accumulation pattern of this strain, as well as lipid fatty acid composition (Table 1), presented similarities to those described above for strains A6 and A11. Also, it seems that this strain is unable to provoke removal of phenolic compounds of OMW neither decolorization of the medium (data not shown). 3.3. Ethanol production The discovery of renewable energy sources is nowadays a very important priority, with ethanol being one of the most important biofuels produced worldwide. In fact, given that the utilization of the so-called “1st generation” bioethanol, deriving from fermentation of hydrolyzed corn starch (USA) or sucrose juice (Brazil), confronts on the food vs fuel dilemma (Koutinas et al., 2014; Pimentel et al., 2007), the production of the so-called “2nd generation” bioethanol (deriving through valorization of several waste streams) is considered as a very important scientific priority worldwide (Sarris and Papanikolaou, 2016). Production cost of the 1st or 2nd generation ethanol should be competitive to the that of gasoline (Chandel et al., 2007). However, in many cases, especially concerning ethanol production using agro-industrial residues as feedstocks, the absence of subsidies would reduce or even cease this production (Pimentel et al., 2007). In the above-mentioned report, it had been considered that if the production costs of a L of ethanol were added to the tax subsidy cost, then the total cost per L would range between 0.79 and 1.21 USD. On the other hand, in several cases in which more complex substrates than simple sugars are implicated in the process (e.g. starch, cellulose), pretreatment phases are required (Sarris and Papanikolaou, 2016) and the more the number processes and unit operations implicated into the bioconversion increases, the more the final ethanol cost is higher (Azmi et al., 2012). For instance, in countries like Brazil, sugarcane (or sucrose-rich juices) is preferentially used as substrate instead of starch (e.g. from corn grain) since they are more efficient feedstocks, resulting in a production cost ranging between 0.22 and 0.33 USD per L (Pimentel et al., 2007). In any case, a significant part of the bioethanol fermentation cost (i.e. 40e60%) refers to the substrate cost (Azmi et al., 2012) suggesting that choice of a low- or even negative-cost material amenable to be converted into ethanol would significantly reduce the whole process economy (Chandel et al., 2007). In recent reports in which various fermentation configurations would have been envisaged [i.e. separate hydrolysis and fermentation (Sarris and Papanikolaou, 2016)] the cost of corn-derived ethanol was found to be lower than 1.0 USD per L, often ranging between 0.25 and 0.50 USD per L (Chandel et al., 2007). Benvenga et al. (2016) reported that for the period 2002e2013, the alcohol production cost from cassava root, a starch-rich residue, was found to vary from 0.04 to 0.62 USD per L. The carbon substrate choice is, therefore, of primordial importance for the conversion of biomass into ethanol, since it represents an important cost of the whole process (Azmi et al., 2012; Chandel

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Fig. 5. Kinetics of growth, lipid accumulation and substrate consumption of Yarrowia lipolytica LGAM S (7) cultivated in flasks on media containing OMW. Culture conditions and abbreviations: as in Fig. 2.

et al., 2007). Besides, an important issue that should be taken into consideration refers to the fact that significant quantities of wastewaters are generated during bioethanol production. For instance, Pimentel et al. (2007) reported that 12 L of wastewaters are released per L of 99.5% purity ethanol produced. However, this parameter has not been taken into consideration in most of the previously studies concerning ethanol production cost. Under this point of view, in the present investigation, the conversion of OMW into ethanol by using yeast strains was proposed. Indeed, since the OMW employed here was not rich in sugars, blends with crude glucose were utilized, the OMW being considered as simultaneous substrate and process water, increasing the economic efficiency of the process. It is evident that a more economically viable scheme would have been proposed if instead of commercial glucose, a concentrated sugar-containing wastewater diluted with OMW would have been used (Sarris and Papanikolaou, 2016). Concerning the biological restrictions imposed by OMW used as substrate for bioethanol production, it is evident that the microorganisms implicated in the process should present a significant resistance into the phenolic compounds. Therefore, the strain S. cerevisiae MAK-1, which has been revealed capable to grow on media containing phenolic compounds in high concentrations (Sarris et al., 2014, 2013), was such a choice. On the other hand, in preliminary studies, the microorganism Candida tropicalis LFMB 16 has shown the ability to produce ethanol on glucose based media (data not shown). Moreover, it has been indicated that strains of C. tropicalis present a remarkable resistance against the phenolic compounds found into OMW based media (Ettayebi et al., 2003), while recently, amongst other microbial species employed, it has been demonstrated that C. tropicalis strains can perform aerobic treatment of OMW with simultaneous bioethanol production (Jamai and Ettayebi, 2015). Therefore, both C. tropicalis LFMB 16 and S. cerevisiae MAK-1 were selected as ethanol production candidates and were cultivated on OMW based media under conditions which favored ethanol production. Both strains were cultivated at low pH values (i.e. C. tropicalis at 4.5 and S. cerevisiae at 3.5), since as it has already been mentioned, low pH values can protect cultures from bacterial contaminations and therefore are desirable for large scale applications.

Bioreactor cultures of C. tropicalis LFMB 16 were conducted in OMW enriched with 65 g/L glucose. The presence of OMW in the medium at the appropriate dilution so as to obtain initial phenolic compounds concentration less than 1.5 g/L, gave satisfactory results as regards the maximum production of ethanol (EtOHmax ¼ 21.9 g/ L), since similar results were obtained in the absence of OMW (Table 3a). Moreover, conversion yield of ethanol produced per unit of substrate consumed (YEtOH/S) seemed to be unaffected by the presence of OMW in the growth medium, confirming that ethanol synthesis was not inhibited by OMW components. On the contrary, biomass production was negatively affected, as on OMW plus glucose fermentation Xmax was only 2.6 g/L, while Xmax achieved on glucose fermentation in flask cultures was 7.7 g/L (data not shown). Concerning decolorization of the culture medium, color removal was achieved at a ratio of 56e58%, contrariwise the maximum removal of phenolic compounds was only 16.5%, demonstrating that phenolic removal does not result a corresponding reduction in color. In media containing OMW and 65 g/L glucose S. cerevisiae MAK1 produced 20.6 g/L ethanol that is higher to the production achieved in the absence of OMW (Table 3b). However, the YEtOH/S value was lower in the presence of OMW (i.e. 0.34 g/g vs 0.45 g/g observed in the absence of OMW). In addition, the Xmax obtained in OMW enriched with glucose medium was 11.5 g/L (YX/S ¼ 0.18 g/g), while when glucose was used as sole carbon source only Xmax ¼ 4.8 g/L (YX/S ¼ 0.18 g/g) were produced (data not shown). Increased growth, but to a lesser extent, was also observed in the presence of OMW in bioreactor cultures. It seems therefore that several nutrients found in OMW stimulate growth of S. cerevisiae. Contrariwise, EtOHmax production was lower on OMW enriched with glucose media, as 31.3 g/L were produced, against 34.0 g/L that were produced in the absence of OMW, but the YEtOH/S values were similar (around 0.45 g/g). Finally, comparing flasks and bioreactor cultures in the presence of OMW, higher EtOHmax production was achieved during fermentation in bioreactor, probably due to the lower dissolved oxygen concentration achieved in bioreactor. Ethanol yields of S. cerevisiae ranged between 0.34 and 0.49 g/g, and this was depended on the initial concentration of sugar and the initial C/N ratio into the medium (Çaylak and Vardar Sukan, 1998;

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Fig. 6. Kinetics of growth and lipid accumulation (a) and citric acid synthesis and substrate consumption (b) of Yarrowia lipolytica LGAM S (7) cultivated in flasks on media containing OMW enriched with glycerol 50 g/L. Culture conditions and abbreviations: as in Fig. 4; C.A., citric acid.

Table 3 Growth, consumed reducing sugars, phenolic removal and decolorization at the fermentation time in which maximum ethanol production was achieved by Candida tropicalis LFMB 16 (a) and Saccharomyces cerevisiae MAK-1 (b) cultivated in flask or in bioreactor cultures on media containing OMW enriched with glucose or sole glucose. Strain

Medium

Type of culture

t (h)

EtOHmax (g/L)

(a)

C. tropicalis LFMB 16

(b)

S. cerevisiae MAK-1

OMW þ Glucose 65 g/L Glucose 65 g/L Glucose 65 g/L OMW þ Glucose 65 g/L Glucose 65 g/L OMW þ Glucose 65 g/L Glucose 65 g/L

B.C. B.C. F.C. F.C. F.C. B.C. B.C.

46 44 36 20 16 16 17

21.9 22.7 19.5 20.6 18.6 31.3 34.2

± ± ± ± ± ± ±

0.9 0.7 0.6 1.1 1.3 0.9 2.3

X (g/L) 2.6 3.0 7.7 7.9 3.3 5.8 4.5

± ± ± ± ± ± ±

0.02 0.01 0.02 0.4 0.6 0.6 0.5

Consumed R.S. (g/L) 58.9 60.7 59.6 63.6 41.1 72.1 73.5

± ± ± ± ± ± ±

0.9 0.6 0.1 0.8 0.5 0.0 2.8

Phenolic removal (%w/w)

Decolorization (%)

16.5 ± 0.8 e e 15.3 ± 0.9 e 12.9 ± 0.4 e

55.5 ± 0.3 e e 37.4 ± 0.4 e ND e

Culture conditions: Non-aseptic; T ¼ 28  C (a) C. tropicalis LFMB 16: pH 4.5 ± 0.03; initial phenolic compounds 1.2 g/L, (b) S. cerevisiae MAK-1: pH 3.5 ± 0.03; initial phenolic compounds 1.1 g/L. Abbreviations: X, dry biomass; EtOH, ethanol; R.S., reducing sugars; B.C.; bioreactor culture; F.C., flask culture; ND, non-determined.

Hagman et al., 2013; Sarris and Papanikolaou, 2016; Sarris et al., 2009; Wang et al., 2007). Higher conversion of glucose into ethanol than in the current study has been reported by Hagman

et al. (2013) for S. weihenstephan (YEtOH/S ¼ 0.50 g/g), while similar with the current investigation YEtOH/S values have been reported for S. paradoxus, S. uvarum and Kazachstania exiguous

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(Hagman et al., 2013). As far as the maximum quantity of ethanol achieved in the current submission, it is evidently lower than the maximum value achieved in fermentations performed with simple sugars employed as substrates by several S. cerevisiae strains (Çaylak and Vardar Sukan, 1998; Sarris et al., 2009), but it is comparable with the values reported during trials on pretreated OMW (Jamai and Ettayebi, 2015; Massadeh and Modallal, 2008; Zanichelli et al., 2007). For instance, employing conventional substrates (i.e. glucose, starch hydrolysates or sucrose), maximum ethanol yields ranging between 75 and 147 g/L have been reported (Alfenore et al., 2004; Çaylak and Vardar Sukan, 1998; Wang et al., 2007), which are clearly higher than those reported in the present study. Even when cellulose-based materials are employed (e.g. sorghum stalks, bagasses, etc.), higher ethanol yields than those achieved in the current investigation [e.g. ranging between 42 and 63 g/L (Matsakas and Christakopoulos, 2013a, 2013b)] have been reported. However, in the fermentation trials performed in the presence of OMW, the substrate was completely consumed approximately 20 h after the begging of fermentations. Regarding the phenolic removal and decolorization, S. cerevisiae MAK-1 exhibited better decolorization in bioreactor cultures comparing with flasks cultures (i.e. 41.4% and 38.5%, respectively), while the opposite trend was observed for phenolic removal (12.9% against 17.2%). Sarris et al. (2014) investigating the impact of aeration and sterilization on ethanol production by S. cerevisiae MAK-1 reported that under non-aerated conditions ethanol production slightly increased. In the current investigation, it is clear that both C. tropicalis and S. cerevisiae were able to produce ethanol in high quantities, cultivated on OMW based media, under non-aseptic and non-aerated conditions. In addition, removal of phenolics, probably due to the absorbance of phenolics on yeast cell surface, was around 17% for both strains. It is noted that neither polymerization following by precipitation of phenolics, as in the case of white-rot fungi (D'Annibale et al., 1998), nor phenolics degradation, as in molds such as Aspergillus niger (Hamdi et al., 1992) have been observed. Color removal was efficiently performed by both strains, although results were better for C. tropicalis LFMB 16 (maximum decolorization at a ratio of 57.7% by C. tropicalis was achieved, against 41.4% by S. cerevisiae). Although the nitrogen concentration may favor yeast proliferation, thus indirectly increase phenolics removal, in the experiments performed during this research no significant effect of yeast extract concentration on phenolics removal has been observed. 4. Conclusions OMW could successfully be used as substrate for SCO, mannitol, citric acid and ethanol production by selected yeast strains of L. starkeyi, Y. lipolytica, C. tropicalis and S. cerevisiae. Because of the inhibitory effect of phenolics on the microbial growth, a suitable dilution of OMW is needed, as well as an enrichment of the medium with carbon sources, such as glucose and glycerol. OMW enriched with glycerol, firstly used in this paper, maybe an important medium for citric acid and/or mannitol production. Utilization of OMW as feedstock for microbial metabolites production can decrease the production cost of the target product and diminish environmental pollution caused by OMW. Actually, an effective removal of phenolic compound from OMW was performed, while some strains (i.e. C. tropicalis and S. cerevisiae) were able to provoke medium decolorization. Acknowledgements Financial support for this study has been provided by a) Bilateral project Greece-Hungary 2012-1014 entitled “Microbial conversions

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